PHYSIOLOGICAL ADAPTATION TO NUTRIENT LIMITATION IN A MARINE OLIGOTROPHIC ULTRAMICROBACTERIUM Sphingopyxis alaskensis
MARTIN OSTROWSKI
A thesis submitted in fulfilment of the requirements for the degree of Doctor of Philosophy
School of Biotechnology and Biomolecular Sciences Faculty of Science The University of New South Wales, Australia
October 2006 UNIVERSITY OF NEW SOUTH WALES Thesis/Project Report Sheet
Surname or Family name: OSTROWSKI First name: MARTIN Other name/s: - LUKE Abbreviation for degree as given in the University calendar: PhD School:BIOTECHNOLOGY AND BIOMOLECULAR SCIENCES Faculty: SCIENCE Title: PHYSIOLOGICAL ADAPTATION TO NUTRIENT LIMITATION IN A MARINE OLIGOTROPHIC ULTRAMICROBACTERIUM Sphingopyxis alaskensis
Abstract 350 words maximum:
Sphingopyxis (formerly Sphingomonas) alaskensis, a numerically abundant species isolated from Alaskan waters and the North Sea represents one of the only pure cultures of a typical oligotrophic ultramicrobacterium isolated from the marine environment. In this study, physiological and molecular characterization of an extinction dilution isolate from the North Pacific indicate that it is a strain of Sphingopyxis alaskenis, extending the known geographical distribution of this strain and affirming its importance as a model marine oligotroph. Given the importance of open ocean systems in climatic processes, it is clearly important to understand the physiology and underlying molecular biology of abundant species, such as S. alaskensis, and to define their role in biogeochemical processes.
S. alaskensis is thought to proliferate by growing slowly on limited concentrations of substrates thereby avoiding outright starvation. In order to mimic environmental conditions chemostat culture was used to study the physiology of this model oligotroph in response to slow growth and nutrient limitation. It was found that the extent of nutrient limitation and starvation has fundamentally different consequences for the physiology of oligotrophic ultramicrobacteria compared with well-studied copiotrophic bacteria (Vibrio angustum S14 and Escherichia coli). For example, growth rate played a critical role in hydrogen peroxide resistance of S. alaskensis with slowly growing cells being 10, 000 times more resistant than fast growing cells. In contrast, the responses of V. angustum and E. coli to nutrient availability differed in that starved cells were more resistant than growing cells, regardless of growth rate.
In order to examine molecular basis of the response to general nutrient limitation, starvation and oxidative stress in S. alaskensis we used proteomics to define differences in protein profiles of chemostat-grown cultures at various levels of nutrient limitation. High-resolution two-dimensional electrophoresis (2DE) methods were developed and 2DE protein maps were used to define proteins regulated by the level of nutrient limitation. A number of these proteins were identified with the aid of mass spectrometry and cross-species database matching. The identified proteins are involved in fundamental cellular processes including protein synthesis, protein folding, energy generation and electron transport, providing an important step in discovering the molecular basis of oligotrophy in this model organism.
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Table of Contents I
List of Figures V
List of Tables VII
List of Abbreviations VIII
Certificate of Originality X
Acknowledgements XI
Publications arising from this work XII
ABSTRACT XIV 1 GENERAL INTRODUCTION ...... 1 1.1 INTRODUCTION ...... 2 1.2 OLIGOTROPHIC ENVIRONMENTS ...... 3 1.2.1 General observations...... 3 1.2.1 An overview of the marine microbial community ...... 4 1.2.2 Factors affecting microbial growth...... 8 1.3 DISTINCTIONS AND DEFINITIONS ...... 12 1.3.1 The size of small organisms...... 13 1.3.2 Copiotrophs vs Oligotrophs...... 14 1.4 OLIGOTROPHIC ISOLATES ...... 16 1.4.1 Predicted properties of oligotrophs ...... 16 1.4.2 The extinction dilution method for isolating oligotrophs ...... 17 1.5 DESCRIPTION OF SPHINGOMONAS ALASKENSIS RB2256...... 21 1.5.1 Isolation of S. alaskensis...... 21 1.5.2 General growth characteristics ...... 23 1.5.3 Starvation and stress resistance...... 25 1.6 HYPOTHESIS ...... 26 1.6.1 Overall objectives...... 27 1.6.2 Specific aims ...... 27 2 MOLECULAR AND CHEMOTAXONOMIC CHARACTERISATION OF AN ABUNDANT ULTRAMICROBACTERIUM ISOLATED FROM THE OLIGOTROPHIC NORTH PACIFIC...... 29 2.1 BACKGROUND ...... 30 2.2 MATERIALS AND METHODS...... 32 2.2.1 Bacterial strains and culture conditions ...... 32 2.2.2 Alcohol precipitation...... 32 2.2.3 Estimation of nucleic acid concentration ...... 32 2.2.4 Restriction enzyme digestion...... 33 2.2.5 Agarose gel electrophoresis and visualisation ...... 33 2.2.6 Genomic DNA extraction...... 34 2.2.7 Amplification of 16S rDNA sequences ...... 34 2.2.8 Purification and nucleotide sequencing of amplification products ...... 35 2.2.9 Phylogenetic analysis of 16S rDNA gene sequences...... 35 2.2.10 DNA-DNA hybridisation and mol%G+C content analysis ...... 35 2.2.11 Fatty acid analysis...... 36 2.3 RESULTS...... 38 2.3.1 Sequencing and phylogeny of strain AFO1 ...... 38 2.3.2 DNA base composition and DNA-DNA hybridisation ...... 42 2.3.3 Fatty acids...... 43 2.4 DISCUSSION...... 45 2.4.1 Ecological implications of S. alaskensis sp. strain AFO1...... 45 2.4.2 Reclassification of Sphingomonas alaskensis to Sphingopyxis alaskensis 47 2.4.3 Conclusion ...... 48 3 CONTINUOUS CULTURE METHODS FOR INVESTIGATING PHYSIOLOGICAL RESPONSES TO NUTRIENT LIMITATION IN MARINE BACTERIA...... 50 3.1 BACKGROUND ...... 51 3.1.1 Small scale chemostats ...... 56 3.1.2 Theoretical considerations: growth rates and nutrient flux...... 57 3.2 MATERIALS AND METHODS...... 59 3.2.1 Bacterial strains, media and culturing conditions...... 59 3.2.2 Total cell counts and morphology...... 60 3.2.3 Viability measurements...... 61 3.3 RESULTS...... 62 3.3.1 Validation of mini chemostats...... 62 3.3.2 Substrate limitation ...... 63 3.3.3 Steady-state growth and viability of S. alaskensis and V. angustum S14 in chemostat culture...... 64 3.3.4 Morphology and cell dimensions ...... 67 3.4 DISCUSSION...... 71 3.4.1 The impact of nutrient limitation on the viability, morphology and cell dimensions of S. alaskensis and V. angustum S14...... 71 3.4.2 Growth yield and maintenance energy of S. alaskensis and V. angustum S14 in response to nutrient limitation...... 74 4 PHYSIOLOGICAL RESPONSES TO NUTRIENT LIMITATION AND STRESS FOR AN OLIGOTROPHIC ULTRAMICROBACTERIUM, SHINGOPYXIS ALASKENSIS...... 77
4.1 BACKGROUND ...... 78 4.2 MATERIALS AND METHODS...... 81 4.2.1 Bacterial strains, maintenance and culturing conditions...... 81 4.2.2 Viability measurements...... 81 4.2.3 Stress exposure experiments ...... 82 4.2.4 Determination of catalase activity ...... 82 4.3 RESULTS...... 84 4.3.1 The effect of growth rate on hydrogen peroxide resistance in S. alaskensis...... 84 4.3.2 The effect of temperature on peroxide resistance ...... 86 4.3.3 The effect of carbon vs nitrogen limitation on peroxide resistance ...... 88 4.3.4 The effect of growth rate on hydrogen peroxide resistance in V. angustum S14...... 90 4.3.5 The effect of growth rate on hydrogen peroxide resistance in E. coli....92 4.3.6 Catalase activity and hydrogen peroxide resistance in S. alaskensis....94 4.3.7 The effect of growth rate on resistance to ultraviolet radiation in S. alaskensis ...... 97 4.3.8 The effect of growth rate on heat stress and freeze-thaw resistance in S. alaskensis ...... 98 4.3.9 The possible role of cellular pigments in stress resistance of S. alaskensis ...... 103 4.3.10 The effect of growth conditions and nutrient limitation on the abundance of nostoxanthin of S. alaskensis ...... 104 4.4 DISCUSSION...... 107 4.4.1 Growth rate control of hydrogen peroxide resistance ...... 107 4.4.2 Possible mechanisms of growth rate control of hydrogen peroxide resistance...... 109 4.4.3 Resistance to ultraviolet radiation ...... 112 4.4.4 Starvation vs low growth rate induction of peroxide stress resistance 113 4.4.5 Possible role of cell membrane and pigments in overall stress resistance 114 5 AN ASSESSMENT OF TWO DIMENSIONAL PROTEIN REFERENCE MAPS FROM S. ALASKENSIS RB2256 ...... 118 5.1 BACKGROUND ...... 119 5.2 MATERIALS AND METHODS...... 122 5.2.1 Biomass sampling and preservation...... 122 5.2.2 Pulse labelling of chemostat cultures...... 122 5.2.3 Sample preparation ...... 122 5.2.4 Isoelectric focusing...... 123 5.2.5 Equilibration and SDS-PAGE...... 123 5.2.6 Gel staining...... 124 5.2.7 Image acquisition and spot detection...... 124 5.3 RESULTS AND DISCUSSION ...... 127 5.3.1 Characteristics of silver stained and radiolabelled 2DE images ...... 127 5.3.2 Comparison of silver stained and radiolabelled 2DE images...... 130 5.3.3 Possible explanations for quantitative differences between methods ..131 5.3.4 Representation of the S. alaskensis proteome in the pH 4-7, Mr 10-200 kDa window...... 135 5.3.5 Conclusion ...... 139 6 EXAMINATION OF GENE EXPRESSION IN RESPONSE TO NUTRIENT LIMITATION IN S. ALASKENSIS...... 140 6.1 BACKGROUND ...... 141 6.2 MATERIALS AND METHODS ...... 146 6.2.1 Two-dimensional electrophoresis, image acquisition and analysis.....146 6.2.2 Preparative 2DE and semi-dry blotting ...... 146 6.2.3 Blot staining...... 146 6.2.4 Sample preparation and in-gel trypsin digestion...... 147 6.2.5 Mass spectrometry analysis ...... 148 6.2.6 Amino acid and Edman degradation...... 149 6.3 RESULTS...... 150 6.3.1 Differential expression of proteins observed on silver stained gels ....150 6.3.2 Differential expression of pulse labelled proteins ...... 155 6.3.3 Identification of protein spots on the basis of peptide mass fingerprinting and Edman sequencing...... 160 6.3.4 Identification of protein spots by LC-ESI MS/MS...... 162 6.4 DISCUSSION...... 165 6.4.1 Differential expression of proteins in steady state chemostats...... 165 6.4.2 Cross species identification ...... 170 6.4.3 Identified proteins ...... 172 7 GENERAL DISCUSSION...... 176 7.1 SUMMARY ...... 177 7.2 EXTINCTION DILUTION, ...... 178 7.2.1 S. Alaskensis sp. strain AFO1...... 178 7.2.2 Extinction dilution and lab domestication...... 179 7.3 THE IMPACT OF NUTRIENT LIMITATION ON THE PHYSIOLOGY OF MARINE BACTERIA...... 182 7.3.1 Distinct life strategies?...... 182 7.3.2 Comparison of S. alaskensis with Peligibacter ubique ...... 184 7.4 THE MECHANISMS OF STRESS RESISTANCE IN S. ALASKENSIS ...... 186 7.4.1 Carotenoids...... 188 7.5 GENE EXPRESSION DURING NUTRIENT LIMITED GROWTH AND STARVATION ...... 189 7.6 FUTURE PROSPECTS...... 193 7.6.1 Molecular genetics ...... 193 7.6.2 Cross species identification of proteins: environmental proteomics ...194 7.7 CONCLUDING REMARKS ...... 195 8 REFERENCES ...... 197 LIST OF FIGURES
Figure 2.1 Distance-matrix tree of selected Sphingomonas 16S rDNA 41 sequences
Figure 3.1 Schematic diagram of the chemostat apparatus used in this study 55
Figure 3.2 Development of total cell numbers and CFU in a glucose limited 66 chemostat of S. alaskensis
Figure 3.3 Molar growth yield and cell numbers of S. alaskensis and V. 67 angustum S14 grown in glucose limited chemostats at different specific rates of growth
Figure 4.1 Percent survival of S. alaskensis grown at different specific growth 85 rates in glucose-limited chemostats after exposure to 25mM hydrogen peroxide for 60 min
Figure 4.2 Percent survival of nutrient limited chemostat grown S. alaskensis 87 cells following exposure to 25 mM hydrogen peroxide for up to 60 min
Figure 4.3 Percent survival of S. alaskensis in response to the concentration of 89 hydrogen peroxide
Figure 4.4 Percent survival of nutrient limited chemostat grown S. alaskensis 91 cells following exposure to 25 mM hydrogen peroxide for up to 60 min
Figure 4.5 Percent survival of V. angustum S14 following exposure to 2mM 93 hydrogen peroxide for up to 60 min after growth at various rates in glucose limited chemostats
Figure 4.6 Percent survival of log-phase, starved and chemostat grown E. coli 95 K12 UNSW002900 after exposure to 25 mM hydrogen peroxide for up to 60 min
Figure 4.7 Survival of batch grown S. alaskensis cells following exposure to 99 UV radiation. UV doses ranged from 0 to 10,000 J m-2
Figure 4.8 Percent survival of batch and chemostat grown S. alaskensis 101 following exposure to 4000 Jm-2 ultraviolet radiation
Figure 4.9 Percent survival of exponentially growing S. alaskensis following 102 heat stress
Figure 4.10 Absorbance spectra of cellular methanol extracts of S. alaskensis 105
Figure 4.11 Molecular structure of nostoxanthin 105
Figure 4.12 Relative abundance of carotenoids extracted from S. alaskensis 106 grown in glucose-limited chemostats and batch cultures
35 Figure 5.1 The incorporation of [ S] L-methionine for cultures grown at 129 35 Figure 5.1 The incorporation of [ S] L-methionine for cultures grown at 129 different glucose limited rates in chemostat cultures
Figure 5.2 Comparison of protein gel images prepared from 2DE profiles of 134 silver-stained total protein and radioactively labelled proteins of S. alaskesnis grown in chemostat culture
Figure 5.3 Distinct features present in radiolabelled and silver stained gel 135 images
Figure 5.4 Theoretical 2DE protein maps for -proteobacteria closely related 138 to S. alaskensis
Figure 6.1 Outline of protein identification strategy on the basis of micro- 145 Liquid Chromatography Electrospray Ionisation Tandem Mass Spectrometry ( LC-ESI MS/MS)
Figure 6.2 A comparison of silver-stained 2DE gel images of total protein 152 taken from S. alaskensis grown in glucose limited chemostats
Figure 6.3 Differential intensity for silver stained protein spots from S. 154 alaskensis cultures grown at low relative to high rates of growth in glucose-limited chemostats
Figure 6.4 A comparison of 2DE gel images of protein from S. alaskensis 157 grown in glucose limited chemostats
Figure 6.5 Differential intensity for radiolabeled protein spots from S. 159 alaskensis cultures grown at low relative to high rates of growth in glucose-limited chemostats
Figure 6.6 Two dimensional reference image showing the protein spots 169 characterised and their regulation grouping LIST OF TABLES
Table 1.1 Description of terms relating to the small size of bacteria 16
Table 1.2 General characteristics of selected marine oligotrophic isolates 20
Table 2.1. Changes and gaps relative to S. alaskensis RB2256T 16S rDNA 40 sequence
Table 2.2 DNA base composition and DNA:DNA hybridisation values 42 between S. alaskensis RB2256, strains AFO1and KT-1, and S. macrogoltabidus
Table 2.3 FAME analysis of fatty acids from S. alaskensis RB2256 and 44 Sphingomonas strains AFO1 and KT-1
Table 3.1 Assumed physiological constants and their relationships during 56 steady state cultivation in chemostat culture
Table 3.2 Mixing time, a measure of homogeneity in ‘classic’ large scale and 63 small scale chemostat culture vessels
Table 3.3 Size and volume measurements for S. alaskensis determined by 70 transmission electron Microscopy and phase contrast microscopy
Table 3.4 Size and volume measurements for V. angustum S14 determined 70 from estimates based on phase contrast microscopy
Table 4.1 Catalase activity measured in whole cells of S. alaskensis grown in 98 glucose-limited chemostats and batch cultures
Table 5.1 A comparison of the features of 2DE reference maps generated by 130 silver staining of total protein and pulse labelling of newly synthesised proteins with radioactive methionine
Table 6.1 Examples of the number of unique, differentially expressed and total 143 proteins analysed in selected bacterial 2DE projects
Table 6.2 Summary of the number of features detected and the number of spots 151 analysed from replicate silver-stained and radiolabelled gel images
Table 6.3 Summary of characteristics for differentially expressed protein spots 153 on silver stained 2DE gels
Table 6.4 Summary of characteristics for differentially expressed protein spots 158 on silver radiolabeled 2DE gels
Table 6.5 Summary of differentially expressed protein spots on silver stained 160 and radiolabelled 2DE gels
Table 6.6 Proteins characterised by mass spectrometry or Edman sequencing 164 from the S. alaskensis proteome LIST OF ABBREVIATIONS
2DE Two Dimensional gel Electrophoresis A Adenosine ADP adenosine diphosphate APAF AustralianProteomeAnalysis Facility ASW Artificial Sea Water ASW-G Artificial Sea Water-Glucose ATP adenosine triphosphate ATP-ase Adenosine triphosphate synthase BCA Bicinchoninic Acid BCIP 5-bromo-4-chloro-3-indolyl phosphate BLAST Basic Local Alignment Search Tool bp base pair(s) cAMP cyclic Adenosine Mono Phosphate CBB Coomasie Brilliant Blue CFU Colony Forming Units CHAPS 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate Ci Curie CID Collision-Induced Dissociation CRP cAMP Receptor Protein CS-screen Carbon Sulphur-screen CTAB Hexadecyltrimethylammoniumbromide d day(s) Dia. diameter db database DEPC diethyl-pyrocarbonate DFAA dissolved free amino acids DI Differential Intensity DNA Deoxyribonucleic acid dNTP deoxy nucleotide triphosphate DOC Dissolved Organic Carbon dpm disintegration per minute DTT Dithiothreitol EDTA Ethylene diamine tetraacecitc acid, trisodiumsalt ESI-MS Electron Spray Ionization-Mass Spectrometry EtBr Ethidium Bromide FAS Filtered-Autoclaved Seawater FISH Fluorescent In Situ Hybridation g gram(s) GSL glycosphingolipid GTP Guonisine triphosphate h hour(s) HPLC High Performance Liquid Chromatography IEF IsoElectric Focusing IPG Immobilized pH Gradient J Joules kbp kilo basepair(s) kDa kiloDalton(s) kV kilo Volt LB Luria Bertani μLC-ESI MS Liquid Capillary-ElectronSpray IonisationMass Spectrophotometer LMP Low Melting Point MALDI-TOF Matrix Assisted Laser Disorption Ionisation – Time of Flight Mb mega basepairs Min Minutes MNB Marine Nutrient Broth MOWSE Molecular Weight Search MS Mass Spectrometry Mr molecular we ig h t NCBI National Center for Biotechnology Information nm nanometer(s) nrdb non redundant database OD Optical Density
OD433 Optical Density at 433 nm
OD610 Optical Density at 610 nm PAGE PolyAcrylamide Gel Electrophoresis PBS Phosphate buffer saline PCR Polymerase Chain Reaction pI Isoelectric point PMF Peptide Mass Fingerprinting Ppm partspermillion PVDF polyvinylidine difluoride RNA Ribonucleic acid rpm revolutions per minute SDS sodium dodecyl sulphate s second(s) sp. species TAE Tris-Acetic acid-EDTA TCA Trichloroacetic acid TE Tris-EDTA TEMED N,N,N’,N’-Tetramethylethylene-diamine TOF Time of Flight UV-B Ultra Violet –B (wavelengths, 290-320 nm) Vh volt-hours CERTIFICATE OF ORIGINALITY
I hereby declare that this submission is my own work and to the best of my knowledge it contains no material previously published or written by another person, nor material which to a substantial extent has been accepted for the award of any other degree or diploma at UNSW or any other education institution, except where due acknowledgement is made in the thesis. Any contribution made to the research by others, with whom I have worked in UNSW or elsewhere, is explicitly acknowledged in the thesis.
I also declare that the intellectual context of this thesis is the product of my own work, except to the extent that assistance from others in the project’s design and conception and linguistic expression is acknowledged.
Martin Ostrowski ACKNOWLEDGEMENTS
This thesis would not have been possible without the contributions of many people, for which I am deeply indebted. I would like to sincerely thank my supervisor Associate Professor Rick Cavicchioli for his guidance, undersdtanding, patience and constant encouragement. In addition I would also like to thank all of my supervisors, past and present, Jan Gottschal, Tom Stewart, Dave Scanlan and Paul March for fostering my love of science and sharing my successes and failures. Thank you to Dr. Valerie Wasinger and Dr. Garry Corthals, formerly of the Protein Lab at the Garvan Institute, for their assistance and kind advice in performing ESI-MS analysis. To Dr John Bowman for DNA-DNA hybridisation and Dr David Nichols for fatty acid analysis. I must especially acknowledge my colleagues Fitri Fegatella and Amber Goodchild for their insight, cooperative assistance and thoughtful discussions. Thanks to Dr Neil Saunders for teaching me how to use a computer. Thanks to other members of the laboratory, Julie Lim, Torsten Thomas, Nuria, Laura and Sohail for their support and encouragment. Thanks also to Caiyan, Anne-Carlijn, Andrew, Phil, Michael, Kristine and Christine for being such lovely and enthusiastic students. Parts of this work were supported by the Australian Research Council and the Department of Science, Industry and Tourism. My subsistence was supported by an Australian Postgraduate Award, the Australian Society for Microbiology Research Trust Award, the School of Microbiology and Immunology and my supervisor, and this is duly acknowledged. I am truly grateful for all of the great relationships that I’ve had with people throughout the faculty of Life Science and beyond. Thanks to Shaun for too many things to mention here. Special mention to my great friend Charlie, I will truly miss the many times we’ve been out drinking and the many times we’ve shared our thoughts. I wonder if they ever truly noticed the kind of thoughts we’ve got? Thanks to my parents and extended family who have always projected their comforting familial warmth and support from various corners of the World. Lastly, to Sophie for loyalty, loving support and always being there in times of need. PUBLICATIONS ARISING FROM THIS WORK
1. Ostrowski, M., F. Fegatella, V. Wasinger, M. Guilhaus G.L. Corthals, and R. Cavicchioli. 2004. Cross-species identification of proteins from proteome profiles of the marine oligotrophic ultramicrobacterium, Sphingopyxis alaskensis. Proteomics 4:1779-1788.
2. Cavicchioli. R. and M. Ostrowski. 2003. Ultramicrobacteria. Encyclopedia of Life Sciences. Nature Publishing Group, London
3. Cavicchioli R., M. Ostrowski, F. Fegatella, A. Goodchild, N. Guixa- Boixereu. 2003. Life Under Nutrient Limitation in Oligotrophic Marine Environments: an Eco/Physiological Perspective of Sphingopyxis alaskensis (formerly Sphingomonas alaskensis). Microbial Ecology 45:203-217.
4. Ostrowski, M., R. Cavicchioli, M. Blaauw, and J.C. Gottschal. 2001. Specific growth rate plays a critical role in hydrogen peroxide resistance of the marine oligotrophic ultramicrobacterium Sphingomonas alaskensis strain RB2256. Applied and Environmental Microbiology 67: 1292-1299.
5. Eguchi, M., M. Ostrowski,F.Fegatella,J.Bowman,D.Nichols,T.Nishino and R. Cavicchioli. 2001. Sphingomonas alaskensis strain AF01: an abundant oligotrophic ultramicrobacterium form the North Pacific. Applied and Environmental Microbiology 67:4945-4954.
6. Fegatella, F., M. Ostrowski and R. Cavicchioli. 1999. An assessment of protein profiles from the marine oligotrophic ultramicrobacterium, Sphingomonas sp., strain RB2256. Electrophoresis 20: 2094-2098.
7. Cavicchioli, R., F. Fegatella, M. Ostrowski, M. Eguchi and J. Gottschal. 1999. Sphingomonads from marine environments. Journal of Inductrial Microbiology and Biotechnology 23: 268-272. CONFERENCE PROCEEDINGS
1. Eguchi, M., M. Ostrowski, F. Fegatella, J. Bowman, D. Nichols, T. Nishino, and R. Cavicchioli. 2001. Sphingomonas alaskensis strain AF01: an abundant oligotrophic ultramicrobacterium form the North Pacific. Poster. 9th International Society of Microbial Ecology Congress, Amsterdam, Netherlands. August 25-31.
2. Cavicchioli, R., M. Ostrowski, and F. Fegatella. 1999. Microbial physiology of a model oligotrophic marine ultramicrobacterium. 9th International Congress of Bacteriology and Applied Microbiology at the International Union of Microbiological Societies (IUMS), Sydney, August 16-20.
3. Ostrowski, M., R. Cavicchioli, M. Blaauw, and J.C. Gottschal. 2001. Growth-rate control of hydrogen peroxide resistance of the marine oligotrophic ultramicrobacterium Sphingomonas alaskensis strain RB2256. 9th International Congress of Bacteriology and Applied Microbiology at the International Union of Microbiological Societies (IUMS), Sydney, August 16-20. ABSTRACT
Sphingopyxis (formerly Sphingomonas) alaskensis, a numerically abundant species isolated from Alaskan waters and the North Sea, represents one of very few pure cultures representative of oligotrophic ultramicrobacteria isolated from the marine environment. In this study, physiological and molecular characterization of an extinction dilution isolate from the North Pacific indicate that it is a strain of Sphingopyxis alaskenis, extending the known geographical distribution of this strain and affirming its importance as a model marine oligotroph. Given the importance of open ocean systems in climatic processes, it is clearly important to understand the physiology and underlying molecular biology of abundant species, such as S. alaskensis, and to define their role in biogeochemical processes.
S. alaskensis is thought to proliferate by growing slowly on limited concentrations of substrates thereby avoiding outright starvation. In order to mimic environmental conditions chemostat culture was used to study the physiology of this model oligotroph in response to slow growth and nutrient limitation. It was found that the extent of nutrient limitation and starvation has fundamentally different consequences for the physiology of oligotrophic ultramicrobacteria compared with well-studied copiotrophic bacteria (Vibrio angustum S14 and Escherichia coli). For example, growth rate played a critical role in hydrogen peroxide resistance of S. alaskensis with slowly growing cells being 10, 000 times more resistant than fast growing cells. In contrast, the responses of V. angustum and E. coli to nutrient availability differed in that starved cells being more resistant than growing cells, regardless of growth rate.
In order to examine the molecular basis of the response to general nutrient limitation, starvation and oxidative stress in S. alaskensis we used proteomics to define differences in protein profiles of chemostat-grown cultures at various levels of nutrient limitation. High-resolution two-dimensional electrophoresis (2DE) methods were developed and 2DE protein maps were used to define proteins regulated by the level of nutrient limitation. A number of these proteins were identified with the aid of mass spectrometry and cross-species database matching. The identified proteins are involved in fundamental cellular processes including protein synthesis, protein folding, energy generation and electron transport, providing an important step in discovering the molecular basis of oligotrophy in this model organism. Chapter 1
1GENERAL INTRODUCTION
1 Chapter 1
1.1 INTRODUCTION
Oceans have the highest cellular production rate of any ecosystem on the planet and yet are vast nutrient-limited environments. The high level of productivity is largely due to the phototrophic prokaryotic primary-producers, and the heterotrophic prokaryotes which effect nutrient transformation and remineralisation. The fixation of carbon, nitrogen and phosphorus by marine bacteria, and their subsequent conversion into particulate matter are critically important processes in marine environments that form the basis for the grazing and sinking food chains of the oceans. Heterotrophic ultramicrobacteria are major contributors to oceanic and terrestrial biogeochemical cycles (Whitman et al., 1998). As reservoirs of nutrients in oligotrophic marine ecosystems, they interact with all trophic levels and control the nutrient fluxes via mineralisation thus impacting on the productivity of all marine life. With predictions of increasing ocean oligotrophy as a consequence of global warming (Matear et al., 1999, Wignal et al., 1996) it is clearly important to understand the physiology of this class of bacteria in order to determine the impact they have on oceanic primary production.
A major portion of terrestrial and marine environments are oligotrophic (nutrient poor) and thus, the day-to-day situation for almost all bacteria is that they are limited in their supply of one or more essential nutrients. Despite low levels of nutrients the pelagic marine environment is dominated, in terms of biomass and activity, by small bacteria, variously referred to as ultramicrobacteria, microcells, nanoplankton or picoplankton depending on the terminology adopted by microbial ecologists and physiologists at different points in time (discussed below). This phylogenetically
2 Chapter 1
diverse bacterial size class is believed to proliferate by growing slowly and efficiently competing for limited concentrations of growth nutrients, thereby avoiding outright starvation. The small size, slowly growing bacterial cells that dominate oligotrophic environments are referred to as oligotrophic bacteria, or oligotrophs. Despite the relatively good understanding of the physiology and genetics of readily cultured, faster growing marine bacteria, termed copiotrophs, oligotrophic ultramicrobacteria and the specific roles they play in environmental processes are poorly understood. Knowledge about the physiology of oligotrophs is limited by the availability of environmental isolates, relating largely to the difficulty in culturing these bacteria from their environments and technical limitations for investigating characteristics of marine bacteria under ecologically relevant conditions. To date, most insight into the physiology of the truly low nutrient adapted marine bacteria has been obtained from very few strains, including
Sphingomonas alaskensis isolated by extinction dilution culture as a numerically dominant bacterium from Resurrection Bay, Alaska, and the North Sea (Schut et al.,
1993).
1.2 OLIGOTROPHIC ENVIRONMENTS
1.2.1 General observations
Oligotrophic environments generally lack exogenous supply of nutrients and are defined by a low nutrient flux (< 1 mg carbon per litre per day, Schut et al., 1997) as well as by low absolute concentrations of nutrients (Morita, 1997). According to this definition, a large proportion of the world’s oceans may be considered
3 Chapter 1
oligotrophic (Cole et al., 1988). Despite the low level of nutrients microbial numbers persist on the order of 105 -106 cells ml-1 in the upper 200m of the ocean
(the photic zone) where the bulk of activity occurs. Total prokaryote numbers in the ocean are estimated at 1029 (Whitman et al., 1998), as a result, marine microorganisms contribute a large proportion of the worlds biosphere in terms of carbon, nitrogen and phosphorus. Furthermore, of the three largest microbial habitats (seawater, soil and sediment/soil subsurface), the rates of cellular activity and turnover are highest in the open ocean (Whitman et al., 1998). By virtue of their abundance and biomass heterotrophic prokaryotes in the ocean play an essential role in nutrient transformation and remineralisation. In addition, picophytoplankton (phototrophic prokaryotes and eukaryotes), contribute significantly to global primary production (Li et al., 1992, Campbell et al., 1994, Li,
1994, Vaulot et al., 1995), with estimates as high as 50% of global carbon fixation attributed to this size class (Partensky et al., 1999). Thus, together the smallest heterotrophic and phototrophic cells play an essential role in regulating the accumulation, export, re-mineralisation and transformation of the world’s largest pool of organic carbon (Cole et al., 1988, Carlson et al., 1996) resulting in an ecosystem composed primarily of a microbial food web where prokaryotes and picoeukaryotes represent the most important biological component.
1.2.1 An overview of the marine microbial community
When observed directly, indigenous bacterial communities are rich in carbon and nitrogen and exhibit a low protein and DNA content, they display typical cell volumes in the range from 0.02 – 0.12 m3, around an order of magnitude smaller
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than commonly studied bacteria such as Escherichia coli (Fuhrman, 1981, Simon and Azam, 1989, Schut et al., 1993, Strehl et al., 1999, Button, 2000). Initial attempts to isolate marine bacteria on nutrient rich agar plates revealed a discrepancy of up to three orders of magnitude between plate counts and the observed total number of cells in marine samples (Jannasch and Jones, 1959, Ferguson et al., 1984). Taken together with the observation that a high proportion of early ocean isolates were typically larger (0.34-6.4 m3) and undergo starvation induced miniaturisation processes (Mården et al., 1985, Nyström et al., 1986, Lee and Fuhrman, 1987, Moyer and Morita, 1989a, Schut et al., 1993), it was believed that indigenous microcells represent starved and dormant forms of isolates that could not form colonies on agar plates. It was therefore assumed that in the environment starvation was the natural state of microorganisms. This assertion, however, does not account for a number of observed phenomena listed here. (i) On a per unit volume basis, oceanic microcells exhibit higher activity than the atypically large cells (Douglas et al., 1987, Eguchi and Ishida, 1990, Ouverney and Fuhrman, 1999). (ii) More than 90% of the productivity in pelagic regions is due to free-living, rather than substrate-attached, cells (Cho and Azam, 1988). (iii) Bacteria that remain small when actively growing have been observed and isolated (Ishida and Kadota, 1981, Schut et al., 1993, Rappé et al., 2002). (iv) The global significance and activity of ultramicrobacterial phototrophic cyanobacteria is well established (Partensky et al., 1999), and (v)
Starved, or dormant, bacteria may not become predominant in the ocean while in the non-growing state.
More recently molecular techniques suggest low in situ abundance of typically isolated bacteria (Eilers et al., 2000). Developments with molecular methods
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enabled the relative abundance of specific prokaryotic taxa to be determined without the need to cultivate microorganisms. Initial studies based on SSU rRNA sequence libraries found that the most abundant rDNA sequences obtained did not correspond to cultured species and were distantly related to other rDNA sequences in databases
(reviewed in Giovannoni and Rappé, 2000). These results clearly demonstrated that natural bacterial communities were composed of unknown species that were incapable of forming colonies on commonly used microbiological media. As a consequence, laboratory studies with readily cultured marine bacteria cannot be regarded as a good representation of the numerically dominant indigenous species.
The profound ‘unculturability’ of the microbial community as a whole has severely limited our understanding of the natural state and role of microorganisms from this environment. As a consequence our current knowledge has been largely restricted to direct observation of microbial populations, measurements of in situ activity and the isolation and analysis of community DNA. It is important to note that without detailed characterisation of representative isolates the functional significance of abundant bacterioplankton remains unknown.
Together with rDNA sequencing data from clone libraries the abundance of specific bacterial groups has been estimated using epifluorescence microscopy aided by the development of species-specific probes and fluorescence in situ hybridisation
(Porter and Feig, 1980, and e.g. Glöckner et al., 1999, Cottrell and Kirchman, 2000,
West et al., 2001). A major outcome of molecular based studies is a realisation that the majority of bacterioplantkon belong to just 11 major phylogenetic groups
(Giovannoni and Rappé, 2000). More than 80% of all bacterial rDNA sequences obtained from marine samples belong to only nine phylogenetic groups, most of
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which have no known cultured representatives. Furthermore, uncultured members of the SAR11, SAR116 and Roseobacter lineages of the -Proteobacteria, account for the majority of all rRNA genes that have been identified in seawater while phytoplankton sequences account for about one quarter of the total (Rappé et al.,
2002). Further studies have established distinctions between the composition of coastal and open ocean waters (Fuhrman and Ouverney, 1998, Rappé et al., 2000,
Fuller et al., 2005), particulate-associated and free-living communities (e.g. DeLong et al., 1993) as well as the specific depth distributions of phylogenetic clades
(Moore et al., 1995, West and Scanlan, 2001).
More recently, methods have been used to estimate the relative contribution of actively growing bacteria in the total bacterial pool (Karner and Fuhrman, 1997,
Gasol et al., 1999). An approach recently developed combines microautoradiography and FISH (Lee et al., 1999, Ouverney and Fuhrman, 1999) and allows the uptake of dissolved organic matter (DOM) to be determined in phylogenetically distinct groups. This has been applied to the uptake of amino acids in marine Archaea (Ouverney and Fuhrman, 2000), and supports the observation that different bacterial taxa are responsible for the uptake of low- or high-molecular weight DOM (Cottrell and Kirchman, 2000).
It is important to recognize the limitations of current methods for assessing microbial diversity and activity. Representation in clone libraries is dependent upon methods for extracting community DNA while PCR and probe methods also involve certain biases associated with primer and probe sequences. Enumerating heterotrophic bacteria by microscopy is complicated by the small size of cells 7 Chapter 1
(reviewed in MacGregor, 1999). This causes problems with detection due to the limits of optical resolution, and discriminating heterotrophic cells from auto- fluorescing particles. This also becomes an issue when discriminating small, phototrophic bacteria such as Prochlorococcus spp. which are abundant in oligotrophic waters (Partensky et al., 1999), and can be difficult to distinguish from heterotrophs (Sieraki et al., 1995). Low cellular rRNA content (Fegatella et al.,
1998, Oda et al., 2000) and membrane impermeability also provide limitations for methods relying on FISH or dyes for detection (Pinhassi et al., 1997, Glöckner et al., 1999, Oda et al., 2000). The specific limitations inherent in current methods make it clear that multiple independent approaches are required for understanding the composition and dynamics of the microbial food web.
1.2.2 Factors affecting microbial growth
The fate of prokaryotes in marine food webs has been linked to the grazing activities of nanoplankton organisms (3-20 μm, Sherr and Sherr, 1994, Caron et al., 1999) and to viral lysis (Fuhrman and Suttle, 1993, Furhman, 1999). Consequently, knowledge of the factors that control the abundance, biomass and activity of microorganisms in the open ocean is an essential consideration for the understanding of the biogeochemical fluxes in the ocean (Azam et al. 1984). Despite the fact that the ocean is continuous, it is composed of distinct macro-zones that are affected or defined by currents, stratification, mixing, water depth, photic, non-photic, light intensity, etc. Even within distinct zones, micro-zones persist as a result of microbial distribution, microbial interactions and the colloidal composition of seawater. As a result, microbial populations have evolved a dynamic that reflects the various levels
8 Chapter 1
of heterogeneity within what otherwise appear as a vast homogenous expanse.
Spatial variability includes depth, stratification, aggregates (marine snow) (Alldredge and Silver, 1988), bacteria attached to algae, and even smaller microstructures
(Azam, 1998). Strong inverse gradients of light and nutrients are observed in stratified water columns with intense competition for organic and inorganic nutrients at the surface and higher concentrations of nutrients in light limited areas at the bottom of and below the photic zone (Zubkov et al., 2001, Cavender-Bares et al.,
2001). Temporal variability is also important in some oligotrophic oceans. During the break down of thermoclines in temperate areas (Ducklow et al., 1993, Estrada,
1996, Ducklow, 1999), phytoplankton exhibit higher activities than during either the normal winter or summer periods due to the upwelling of nutrient rich waters to the surface. In tropical latitudes, seasonal variability may also be affected by climatic phenomena such as El Niño (Equatorial Pacific, Ducklow et al., 1995) and Monsoon
(Indian Ocean, Wiebinga et al., 1997). Support for this is illustrated by the distribution of Prochlorococcus spp. in the water column where light intensities vary by up to 4 orders of magnitude (Moore et al., 1995, Partensky et al., 1999, West and
Scanlan, 2001). Throughout this range, physiologically and genetically distinct populations of Prochlorococcus exist which have adapted to high- or low-light intensities (Moore et al., 1998). It has been argued that the co-existence and distribution of distinct eco-types allows the perpetuation of this genus over a wider range of growth conditions than would other wise occur for a single ecotype (Moore et al., 1998). Moreover another illustration of this point is that due to different nutrient acquisition abilities and cell requirements, the growth of co-existing microbial populations can be simultaneously limited by different nutrients, for
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example, phototrophs limited by light and inorganic nutrients and heterotrophs limited by organic carbon (Thingstad and Lignell, 1997).
Not only are large numbers of sub-micron colloids present (Nagata and Kirchman,
1997) but recent experiments demonstrate the spontaneous polymerisation of marine
DOM into polymer gels (Chin et al., 1998). The emerging picture is “seawater as an organic matter continuum, a gel of tangled polymers with embedded strings, sheets, and bundles of fibrils and particles, including living organisms, as hot spots” (Azam
1998). Much higher concentrations of substrates and bacteria can be found in “hot spots” than in the surrounding water column (Pedrós-Alió and Brock, 1983). Pelagic bacteria interact with colloids and particles embedded in the matrix and there is a complex dynamic between DOM and particulate organic matter. These observations go a long way in “fleshing out” the empty space of the ecosystem however, even when we take into account the volume of macro/nano/picoplankton and marine colloids the sum volume of these particles and cells represents less than one hundred thousandth of the total volume of seawater at a cell concentration of 106 ml-1.
Despite the three dimensional matrix and large numbers of particules and cells, the open ocean is an extremely dilute ecosystem composed of mostly ‘empty space’.
1.2.2.1 Nutrient limitation
It is important to consider that the low concentrations of nutrients encountered in oligotrophic environments are in part due to the substrate capture activity of bacteria themselves. Since the open ocean generally lacks an exogenous supply of nutrients, the entire community is ultimately reliant on the release of photoassimilated carbon
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from phytoplankton, nitrogen fixation by diazotrophs in the absence of nitrate/nitrite reductase genes and remineralisation of inorganic nutrients. Taking into account that heterotrophic bacteria dominate the overall biomass of marine ecosystems, by virtue of their surface area and uptake abilities they effectively compete with phytoplankton for mineral nutrients. As a consequence, a close mutual dependence exists between heterotrophic bacteria and phytoplankton (Schut, 1993).
Evidence collected from nutrient addition studies suggest that overall bacterial production is limited by mineral forms of N and P (e.g. Parpais et al., 1996, Sanudo-
Wilhelmy et al., 2002, Thingstad et al., 1998). Specific cases of carbon limitation of heterotrophic bacteria have been found in the equatorial and subarctic Pacific
(Kirchman, 1990, Keil and Kirchman, 1991, Nagata and Kirchman, 1997, Kirchman and Rich, 1997, Church et al., 2000). Aside from methodological problems associated with slow response times of microbial communities (del Giorgio and
Cole, 2000) recent microcosm experiments suggest that the ‘quality’ of added substrates (whether carbon, nitrogen, phosphorus or iron) can have an effect on the efficiency of utilisation of other nutrients, impacting on the observed overall bacterial growth efficiency, productivity and growth rate of communities (del
Giorgio and Cole, 2000). For example, Kirchman (1990) found cases of carbon limitation in the subarctic pacific, however, the best response was found upon addition of amino acids (also a rich source of nitrogen for heterotrophic bacteria,
Keil and Kirchman, 1999). Also, in environments where nitrogen is deficient relative to phosphorus cases have been observed where phytoplankton and the overall bacterial growth rate are limited by phosphorus (Thingstad et al., 1998, Cotner et al.,
1997). It is also possible to design specific probes to assess the physiological state of
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microorganisms in situ. For example, Scanlan et al. (1997) generated antibodies for
PstS, a periplamic binding protein required for high affinity uptake of phophate by marine Synechococcus and Prochlorococcus during phosphate depleted conditions.
Such antibodies have been used to demonstrate seasonal phosphate stress of
Synechococcus and Prochlorococcus populations in the Red Sea (Fuller et al., 2004).
Conflicting results of substrate additions and temporal variability make interpretation of the overall picture difficult. A general picture can be based on the principles outlined above with the growth of heterotrophic bacteria limited by the supply of organic carbon and energy, the growth of phytoplankton limited by the availability of inorganic nutrients and a complex dynamic, in time, space and limiting nutrients, existing between different groups. The observations highlight the need for more specific probes to assess the physiological state of bacteria in situ, a task that cannot be achieved without detailed laboratory study of environmentally relevant isolates.
1.3 DISTINCTIONS AND DEFINITIONS
In a manner reflective of the interdependence of heterotrophic and phototrophic bacteria, microbial ecologists and physiologists are mutually dependent upon each other for the generation and testing of valid ecological hypotheses. Unfortunately, the past decades of research into open ocean ecology were heavily based on phenomenological observations rather than rigorous experimentation under controlled conditions. As a consequence, the terminology adopted by both groups can be polemic, phenomenological and profoundly confusing. Terminology can, however, be extremely useful for creating definitions and making distinctions
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between life strategies. An explanation of currently used terminology is outlined below in an attempt to highlight useful distinctions.
1.3.1 The size of small organisms
The size of microorganisms varies considerably. Even amongst the prokaryotes
(Archaea and Bacteria) cells with volumes of 0.02 – 180,000,000 μm3 have been isolated. At the smallest end of this size spectrum are the ultramicrobacteria. For years microbial ecologists have recognised that they play important roles in the biological cycling of nutrients and in the formation of biomass. Marine ecologists describe a size-graded series of plankton with dimensions ranging from 0.02 μmto
200 m (Sieburth et al., 1978). The terms femto-, pico-, nano- and micro-plankton are used to distinguish size classes with 10-fold increments between 0.02 μmand
200 μm, respectively (Table 1.1). According to these descriptions, ultramicrobacteria most closely correlate with femtoplankton (or femtobacterioplankton). A range of other terms have been coined to describe small microorganisms (Table 1.1). The prefix which has recently been adopted in a number of fields is “nano”; e.g. nan(n)obacteria, nanobe, nanocell and nanosize. The use of “nano” in this context is intended to refer to a size range much smaller (e.g. tens of nanometers) than in the definition of nanoplankton (0.2-2 μm). The use of the term “nanobacteria” may derive from Morita (1988) where it was described as a synonym for ultramicrobacteria.
The term “ultramicrobacteria” was first adopted by Torella and Morita (1981) to describe extremely small bacteria (less than 0.3 μm diameter) isolated from seawater 13 Chapter 1
that formed ‘ultramicrocolonies’ on agar plates, retained their small cell size when growing on agar plates, and grew very slowly in the presence of high concentrations of nutrients. MacDonnel and Hood (1982) modified this description to include isolates from an estuary obtained by filtration through a 0.2 m membrane and which could form normal-sized colonies on low-nutrient agar that were also observed by Hood and MacDonell (1987). In their review, Schut et al., (1997) further modified the description of ultramicrobacteria to include microorganisms which have a cell volume of less than 0.1 μm3, and which retain this volume irrespective of growth conditions. This description using volume as the defining criteria is particularly useful for studies of natural communities as a range of cell shapes are often encountered, and volume provides a measurement of size that is independent of shape. A list of criteria defining ultramicrobacteria are also described by Velimirov (2001).
1.3.2 Copiotrophs vs Oligotrophs
The use of cell size as a defining criteria is also a source of confusion, particularly because cell size can be variable and there are well documented examples where growth rate, growth stage, nutritional status and physiology can effect bacterial cell size and morphology (e.g. Wase and Patel, 1985, Moyer and Morita, 1989a). Perhaps the best studied example is Escherichia coli where the cell mass varies sixfold over a range of growth rates from 0.6 to 2.5 doublings hr-1 (reviewed in Bremer and Dennis,
1996). The increase in cell mass is reflected by an increases in the total amount of protein, RNA and DNA per cell. Despite this well documented example, two functionally distinct types of cells from the environment that display small volumes
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are recognised; ultramicrobacteria and ultramicrocells. Ultramicrocells are formed as a result of starvation induced miniaturisation processes and are characterised by a larger sized (greater than 0.1 μm3) reproductive form and a microcell that can have a volume of less than 0.1 μm3. Unlike ultramicrobacteria which retain a volume of less than 0.1 μm3 when growing, ultramicrocells are dormant, stress-resistant cells, analogous to spores in differentiating bacteria. Accordingly utramicrocells and ultramicrobacteria are products of distinct life strategies adopted by copiotrophic and oligoptrophic bacteria respectively. Copiotrophic bacteria require a high concentration of organic carbon for growth (Poindexter, 1981), they are often associated with nutritionally rich aggregates (DeLong et al., 1993) and are found in relatively higher numbers in coastal regions due to turbulence induced mixing with nutrient rich deeper waters and a considerable input of nutrients from terrestrial sources. When nutrients are scarce starvation induced miniaturisation results in the production of a dormant ultramicrocell (Morita, 1982, 1985, 1988, 1997, Kjelleberg et al., 1993, Holmquist and Kjelleberg, 1993). In contrast, ultramicrobacteria represent the truly low-nutrient adapted oligotrophic species that can become numerically dominant in low nutrient environments (Schut et al., 1997). The presence of these distinct classes of tiny microorganisms is consistent with studies of natural aquatic and soil communities. Microscopic observations of environmental samples reveals cells with volumes of 0.02 – 0.12 m3 (reviewed in Schut et al.,
1997). However, once the environmental samples have been cultured on agar plates, cells typically have volumes on the order of 0.34 - 6.4 m3. Many of the larger cells represent copiotrophs derived from outgrown ultramicrocells, and in practice, are more easily isolated than ultramicrobacteria. Since the isolation of true ultramicrobacteria from marine and soil environments, a key area of research in 15 Chapter 1
contemporary microbial physiology has been to determine what factors affect the culturability of ultramicrobacteria and to compare the physiology of each class.
TABLE 1.1 Description of terms relating to the small size of bacteria
Ultramicrobacteria Microorganism with a cell volume of less than 0.1 m3 that maintains its size with only minor changes, irrespective of growth conditions. Observed by light microscopy. Ultramicrocells Smaller forms (usually starved) of microorganisms that are larger when actively growing. Usually associated with reductive cell division during starvation. Observed by light microscopy. Nan(n)obacteria Possible synonym for UMB. In the literature usually associated with structures in geological samples with sizes ranging from 0.01 – 0.1 μm. Usually associated with uncultured and unsubstantiated descriptions of microorganisms. Observed by electron microscopy. Femtoplankton Marine microorganisms 0.02-0.2 μm Picoplankton Marine microorganisms 0.2-2.0 μm Nanoplankton Marine microorganisms 2.0-20 μm Microplankton Marine microorganisms 20-200 μm Other related Dwarf cells/bacteria, lilliputian cells, femtobacterioplankton, terms miniature cells/bacteria, nanocells, nanosized, nanobe, nano- organisms
1.4 OLIGOTROPHIC ISOLATES
1.4.1 Predicted properties of oligotrophs
According to the vital roles of nutrient uptake and utilisation a list of predicted properties were advanced for a model oligotroph at the Dalhelm Conference (Hirsch et al., 1979). The proposed characteristics include: (i) the possession of high surface per volume ratio (cells are expected to be small or possess prostheca), (ii) preferential usage of metabolic energy for nutrient uptake especially during periods of non-growth, (iii) constitutive uptake nutrient ability, (iv) possession of high
16 Chapter 1
affinity, low-specificity transport systems for simultaneous uptake of mixed substrates, (iv) and the establishment of nutrient reserves following nutrient uptake.
The small size of cells would provide a distinct advantage in terms of grazer avoidance (Morita, 1985) and increased efficiency of nutrient uptake, while nutrient uptake mechanisms were expected to have a broad specificity, be inducible and subject to a minimal amount of catabolite repression in order to ensure simultaneous utilisation of the broadest range of substrates (Poindexter, 1979). Oligotrophs were also expected to regulate their biosynthetic rate in line with nutrient uptake rates
(Poindexter, 1979). Finally, oligotrophs were predicted to have the ability to store diverse nutrients in reserves (Hirsch et al., 1979). Since the proposal of these characteristics a range of physiological studies have been conducted to test their validity (reviewed in Schut et al., 1997). Unfortunately very few of these studies were conducted with oligotrophs, highlighting the need to obtain relevant oligotrophic isolates for laboratory studies.
1.4.2 The extinction dilution method for isolating oligotrophs
The difficulty in isolating oligotrophs from the environment is well documented
(reviewed in Schut et al., 1997). Some of the factors which may restrict the ability to isolate and adapt oligotrophs include: 1. intolerance to high concentrations of nutrients, 2. inappropriate growth substrates, 3. the absence of specific vitamins or growth factors, 4. inhibitory growth substrates or other additives, 5. inactivation by the close proximity to other cells (in colonies on agar plates), 6. susceptibility to the oxidative respiratory burst upon upshift and outgrowth in the presence of fresh nutrients and 7. the deleterious effects of lytic phage.
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To date the extinction dilution method has been the most successful isolation technique. Extinction dilution has been used to obtain numerically abundant strains of S. alaskensis (Button et al., 1993, Schut et al., 1993) and Cycloclasticus oligotrophus (Button et al., 1993, Wang et al., 1996). In the original description of this procedure seawater samples were diluted in filtered and autoclaved natural seawater without additional nutrients until only a few organisms remained in each dilution tube (Button et al., 1993). Isolation without addition of substrates prevented the possibility of substrate toxicity and removed competition for substrates by less abundant indigenous copiotriophs allowing for long term incubation of potentially pure cultures in the highest dilutions. Long-term incubation of these cultures (6-12 months) in the dark and at 5oC initiated an unknown mechanism that enabled the cells to grow on a rich nutrient medium, i.e. a transition from an obligate to a facultatively oligotrophic state (Schut et al., 1993). The nature of this cell transformation is still unclear, however, during subsequent chemostat experiments one of the isolates, RB2256, became unable to form colonies on nutrient rich agar.
This result suggested that the state of adaptation to the external nutrient concentration was an important factor determining whether an oligotrophic isolate would survive on nutrient rich media or not (Schut et al., 1997b). Sensitivity to nutrient concentration has also been demonstrated for SAR11 isolates that are inhibited by dilute proteose peptone (0.001%) (Rappé et al., 2002). The transition mechanism may involve gradual changes in metabolism or cellular composition that allow cells to survive osmotic stress induced by the initial uptake of nutrients and/or the initial oxidative respiratory burst upon outgrowth.
18 Chapter 1
Despite an incomplete understanding of the mechanisms of adaptation, the extinction dilution method has proven to be reproducible. Insights gained from the study of S. alaskensis RB2256 and C. oligotrophus has highlighted the importance of nutrient concentrations and led to modifications of the extinction dilution method, such as the inclusion of vitamins, antioxidants, and a further reduction in the concentration of complex nutrients. Recent applications of the extinction dilution method have resulted in the isolation of previously uncultivated members of the SAR11 clade
(Rappé et al., 2002) as well as novel Gammaproteobacteria from coastal and ocean environments (Cho et al., 2004).
Filtration has also been employed to isolate ultramicrobacteria from the ocean. In contrast to extinction dilution studies however, these attempts have failed (reviewed in Velimirov, 2001). Isolates obtained from the filtrates of 0.2 μm filters have outgrown to larger cells and were therefore likely to be ultramicrocells at the time of filtration. This method however may be useful if the filtrates are initially grown in low-nutrient liquid medium, and then processed in a similar way to those described for the extinction dilution cultures.
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TABLE 1.2. General characteristics of selected marine oligotrophic isolates
Marine isolates Characteristics Sphingomonas Isolated as an abundant species from ocean waters near Alaska alaskensis andintheNorthSea Little variation in cell volume (reviewed in facultatively oligotrophic, obligately oligotrophic upon first Cavicchioli et cultivation al., 2003) High affinity, broad specificity nutrient uptake systems predicted to enable successful competition in oligotrophic waters Simultaneous utilisation of mixed substrates Potential to grow at realistic rates in the ocean Absence of a typical starvation stress response Intrinsically resistant to a range of stresses 3.2 Mb genome Single copy of rRNA operon, maximum 2000 ribosomes cell-1, minimum 200 ribosomes cell-1 Volume = 0.024 m3 Prochlorococcus Most abundant phototrophic organism spp. Global distribution between 40˚N and 40˚S and 0 – 200 m depth Smallest known phototroph (Partensky et al., 1.7-2.4 Mb genome 1999) Accounts for more than 50% of chlorophyll and contributes 30- 80% of the total photosynthesis in the oligotrophic oceans Cycloclasticus Isolated from same location as S. alaskensis from Resurrection oligotrophus Bay, Alaska. Dilute cytoplasm (Button et al., ~3 Mb genome 1998) Utilises only a few aromatic hydrocarbons and acetate as growth substrates Kinetic constants for uptake compatible with growth on ambient concentrations of nutrients in seawater volume = 0.01 m3 HTCC1062 Isolated by extinction dilution from Pacific Waters 27 km off (SAR11) Oregon coast Candidatus Member of the ubiquitous SAR11 lineage based on 16S rDNA Pelagibacter gene sequencing ubique 1.54 Mb genome constant cell volume = 0.01 m3 (Rappé et al., maximum cell density in culture is 3.5 x 106 cells ml-1 2002) 0.001% w/v proteose peptone inhibited growth maximum measured growth rate of 0.58 d-1
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1.5 DESCRIPTION OF SPHINGOMONAS ALASKENSIS RB2256
S. alaskensis RB2256 is the type strain of Sphingomonas alaskensis (Vancanneyt et al., 2001). It is the one of only a few species of marine oligotrophic ultramicrobacteria cultured to date, and is representative of the dominant, pelagic bacterial species in Ressurection Bay, Alaska. S. alaskensis RB2256 is also recognised as one of the earliest oligotrophic isolates and has therefore been the subject of a number of physiological and molecular studies. This strain possesses a number of characteristics that fit the proposed oligotrophic model (Hirsch et al.,
1979) including: (i) a relatively constant ultramicro-size (< 0.1 m3) irrespective of whether it is growing or starved (Schut, PhD Thesis, 1994, Schut et al., 1997a), (ii) relatively slow maximum specific growth rate ( < 0.2 h-1) (Schut et al., 1993, Schut et al., 1995, Eguchi et al., 1996), (iii) the ability to utilize low concentrations of nutrients through a high affinity, broad specificity uptake system (Schut et al., 1995), and also (iv) the ability to take up mixed substrates simultaneously (Schut et al.,
1995, Schut et al., 1997ab).
1.5.1 Isolation of S. alaskensis
S. alaskensis RB2256 and at least 6 other isolates were obtained from a 10 m depth seawater sample from Resurrection Bay, Alaska, serially diluted in filtered aged seawater medium (FAS) in 1990 (Button et al., 1993, Schut et al., 1993). After a period of months bacterial growth was observed by flow cytometry and epifluorescence microscopy at dilutions of 1 x 106 and5x105 and was dominated (>
50% total cells) by small, short rod-shaped bacteria (Schut et al., 1993). Mixed
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cultures with large cells and a variety of different cell types (large rod, cocci, spirilla) were observed to predominate at lower dilutions. The average cell volumes of the small rod-shaped cells were 0.05-0.6 μm3, with the apparent DNA content of 1.0-1.5 fg cell-1. Strain RB2256 and another 8 morphologically similar strains were isolated from serial dilutions demonstrating an initial standing population of 0.2 x 106 cells ml-1 in Resurrection Bay at the time of sampling, indicating that these strains were numerically abundant. These isolates could be subcultured in FAS and oligotrophic medium (2 mg of cassamino acid l-1), but not in high nutrient medium such as full strength marine nutrient agar or broth. Furthermore, growth was never observed on plates, indicating an obligate oligotrophic nature. However, following 6-12 months incubation at 5oC, these isolates gained their ability to form micro colonies on ZoBell
2216E and MPM agars. The acquired ability to grow on richer media represents a transformation from an obligate to facultative oligotroph. In support of the numerical significance of this isolate the presence of this species in Resurrection Bay and the North Sea was also demonstrated by the hybridisation of species specific probes FP1 and FP4 to eight isolates from Ressurection Bay and one isolate (strain
NS1619) from the North Sea (Schut, PhD Thesis, 1994). In addition, the persistence of this species in Resurrection Bay was shown by the detection of cells using
Southern hybridisation of extracted community DNA, more than two years after the initial isolation.
Phylogenetic analysis of strain RB2256 showed that this strain belongs to the -
Proteobacteria lineage, on a single branch within a cluster of Sphingomonas and
Caulobacter (Schut, PhD Thesis, 1994). The genus Sphingomonas was originally described by Yabuuchi et al., in 1990, with the type strain Sphingomonas 22 Chapter 1
paucimobilis but recently split into four new genera by Takeuchi et al. (2001). Prior to 1990, members of the genus Sphingomonas were described as Flavobacterium,
Pseudomonas, Beijerinckia and Arthrobacter. DNA-DNA hybridisation, 16S rDNA sequencing and fatty acid profiles and metabolic characteristics of the remaining isolates indicate that these strains comprise a homogenous genomic species named
Sphingomonas alaskensis with RB2256 as the type strain (Vancanneyt et al., 2001).
At least 7 strains from Resurrection Bay are still available (including RB255,
RB2510, RB2515 and RB2256) from the Culture Collection Laboratorium voor
Microbiologie, University of Gent, Gent, Belgium. To this date, S. alaskensis strains have been isolated from two separate locations: Resurrection Bay Alaska and the
North Sea (Schut et al., 1993), however, the North Sea isolate (strain NS1619) is no longer available.
1.5.2 General growth characteristics
S. alaskensis is an obligately aerobic bacterium that forms opaque yellow, low convex, entire colonies on solid medium (Schut et al., 1993, Vancanneyt et al.,
2001). It has a low specific growth rate (0.13-0.16 h-1 at 23°C) which remains largely unchanged in defined sweater medium with carbon concentrations ranging between 0.8 and 800 mg l-1 (Eguchi et al., 1996). S. alaskensis has the ability to respond rapidly to the addition of excess nutrients. Glucose-limited cultures of S. alaskensis maintain the ability to respond immediately to nutrient up-shift and reach maximum rates of growth without any noticeable lag phase, irrespective of the supplied nutrients (glucose, alanine or acetate) (Eguchi et al., 1996, Fegatella et al.,
1998). The low maximum specific growth rate correlates with a single genome 23 Chapter 1
encoded copy of the ribosomal RNA operon compared to eight or more copies found in the faster growing Vibrio spp. (Fegatella et al., 1998). Even so, S. alaskensis appears to maintain a high cellular concentration of ribosomes and it has been demonstrated that, after periods of outright starvation, a ribosome content 10% of maximum is sufficient to allow cells to immediately respond to nutrient upshift and achieve maximum rates of growth (Fegatella et al., 1998).
S. alaskensis maintains a constitutive high affinity, broad specificity uptake system(s) for amino acids. The alanine uptake system has an affinity for alanine that may exceed any previously reported transport system, is constitutive and is capable of transporting nine other amino acids (Schut et al., 1995). In contrast, the glucose uptake system is inducible and has narrow substrate specificity. Interestingly, alanine and glucose are simultaneously utilised and enable cells to grow at maximum specific growth rates that exceed growth with individual substrates. Both uptake systems use periplasmic binding-proteins which provide an energy dependent transport of substrates against a 105-fold concentration gradient. The uptake systems would enable S. alaskensis to efficiently scavenge and utilise substrates from the environment. Based on the uptake kinetics of these systems, strain RB2256 could grow by using dissolved free amino acids at an in situ doubling time of 12 h to 3 d
(Schut et al., 1995, Schut et al., 1997a), which compares favourably with measured doubling times for bacteria in oligotrophic waters of 5 to 15 d (Fuhrman et al., 1989).
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1.5.3 Starvation and stress resistance
In comparison to the well-documented carbon starvation responses of copiotrophic bacteria, the response in S. alaskensis RB2256 differs in many ways (Schut et al.,
1993, Eguchi et al., 1996, Schut et al., 1997, Fegatella et al., 1998, Fegatella and
Cavicchioli, 2000). The most noteable difference is the absence of reductive cell division which has been described for copiotrophs such as Vibrio ANT 300
(Novitsky and Morita, 1976), V. angustum S14 (Holmquist and Kjelleberg, 1993,
Kjelleberg et al., 1993), Aeromonas, Pseudomonas and Alcaligenes (Moriarty and
Bell, 1993). In addition, starvation does not induce increased protection to other the stress inducing agents (known as cross protection), e.g. hydrogen peroxide (25 mM), ethanol (20%), heat (56˚C) (Eguchi et al., 1996) and UV-B in S. alaskensis
(Joux et al., 1999). Instead, S. alaskensis remains inherently resistant to these stresses. For example the viability of growing cultures decreases less than 2-fold after 30 min exposure to 25 mM hydrogen peroxide (Eguchi et al., 1996). Even though S. alaskensis appears to have no phenotypic response to starvation large changes in ribosome levels (Fegatella et al., 1998), rates of macromolecular synthesis and global gene expression (Fegatella and Cavicchioli, 2000) have been observed, indicating that a specific starvation response exists but that it is significantly different to those of copiotrophic bacteria.
The molecular characteristics of the starvation and outgrowth response have been defined in S. alaskensis (Fegatella, PhD Thesis, 2002). The resumption of growth without lag after periods of starvation and the lack of a transient increase in protein synthesis prior to entry into the starvation phase suggest the lack of a typical
25 Chapter 1
stringent response in this organism (e.g. Flårdh et al., 1994, Cashel et al., 1996).
Like other -proteobacteria, S. alaskensis appears to lack RpoS, perhaps the best characterised starvation response regulator in the literature (Hengge-Aronis, 2002).
In the absence of RpoS, cAMP may serve as a global starvation regulator as protein synthesis levels are matched by the intracellular concentration of cAMP throughout the growth phase. The molecular mechanisms controlling gene expression are likely to be complex and novel.
A great deal of information has been gained by direct comparison of S. alaskensis with well studied copiotrophs, which form the basis of the detailed models of gene regulation and phenotypic responses. However, the relevance of studies on a single isolate in batch culture with high concentrations of nutrients need to be addressed.
In the absence of cell density effects such as quorum sensing and oxygen limitation, studies in batch cultures offer a system in which only the physiology of exponential growth in nutrient replete conditions and the transition to nutrient deplete conditions can be studied. Clearly, in order to understand the physiology of an organism that is believed to proliferate by growing slowly on limited amounts of nutrients we need a better system and more geographically and genetically diverse isolates to study.
1.6 HYPOTHESIS
The model oligotroph Sphingomonas alaskensis possesses novel mechanisms for efficient growth and survival during slow, nutrient limited growth and these mechanisms are distinctly different from those possessed by other well-studied marine copiotrophic bacteria.
26 Chapter 1
1.6.1 Overall objectives
The overall objective of this work is to investigate the physiology of novel oligotrophic isolates under environmentally relevant conditions. Since environmental microorganisms are forced to grow at suboptimal rates as a consequence of the limited availability of growth substrates the aim is to examine the physiology of S. alaskensis during nutrient-limited growth. Chemostats provide a useful tool for examining the effects of chronic nutrient limitaion, although traditional chemostat apparatus is cumbersome and experiments consume vast amounts of media. As an aid to high-throughput analysis of replicates and multiple nutrient limitations, small- scale chemostats with very small working volumes are developed.
Despite a wealth of information describing the responses of copiotrophic bacteria to stress and starvation, very little is known about the impact of nutrient limited growth on the physiology of these organisms. For comparative analyses the physiologies of two well-studied organisms, Escherichia coli and a model marine copiotroph, Vibrio angustum S14 (Nyström and Kjelleberg, 1989, Nyström et al., 1990, 1992,
Holmquist et al., 1993), are also examined under nutrient limitation. Finally, the molecular basis the unique physiology of S. alaskensis will be examined with the aid of proteomics.
1.6.2 Specific aims
27 Chapter 1
1. Chemotaxonomically identify an abundant facultatively-oligotrophic
ultramicrobacterium that was isolated by extinction-dilution culture from the
North Pacific near Toyama Bay, Japan (Chapter 2).
2. Develop small-scale chemostats and methods for analysis of oligotrophic and
copiotrophic bacteria growing under conditions of nutrient limitation
(Chapter 3).
3. Investigate the physiological responses of S. alaskensis to nutrient limited
growth in chemostat cultures and compare the physiology of well-studied
copiotrophic bacteria under the same conditions (Chapter 4).
4. Investigate the effect of nutrient limitation on global gene expression in S.
alaskensis:
a. Establish protocols to generate protein profiles through two-dimensional
gel electrophoresis (2-DE) (Chapter 5).
b. Examine changes in gene expression in carbon limited chemostat cultures:
Perform comparative analysis of cultures growing at different carbon-
limited rates of growth, and identify proteins with altered expression in
each growth condition (Chapter 6).
28 Chapter 2
2MOLECULAR AND CHEMOTAXONOMIC
CHARACTERISATION OF AN ABUNDANT
ULTRAMICROBACTERIUM ISOLATED FROM
THE OLIGOTROPHIC NORTH PACIFIC
29 Chapter 2
2.1 BACKGROUND
A significant focus in marine microbial ecology is the determination of species composition in, as well as the contribution of, each species in biogeochemical cycles.
Molecular studies of microbial diversity and physiological studies with the majority of marine isolates suggest that abundant indigenous marine bacteria are poorly represented in culture collections. Despite advances in assessing species composition and the activity of particular microbial groups in situ, the lack of isolates representing truly low nutrient-adapted microorganisms has limited the understanding of the potential of a large portion of the community.
The extinction dilution method has been used to isolate numerically abundant strains of S. alaskensis and C. oligotrophus from oligotrophic waters in Resurrection Bay,
Alaska (Schut et al., 1993, Button et al., 1993) and representative isolates from the ubiquitous SAR11 and other proteobacterial lineages that have previously defied cultivation efforts (Rappé et al., 2002, Connon and Giovannoni, 2002).
Recently, application of the extinction dilution method by a colleague (Mitsuru
Eguchi) was used to obtain isolates that were abundant in oligotrophic waters near
Japan. After 12 months of serial sub-culturing in filtered autoclaved seawater most of these isolates supported growth in liquid but not on solid medium. However, one isolate, strain AFO1, was able to form colonies on agar plates after 12 months of serial sub culture. Strain AFO1 was isolated from a 10–5 dilution of natural seawater where the microbial count determined by microscopy was 3.1 x 105 cells ml-1
30 Chapter 2
indicating that it was a numerically abundant member of the population at the time of sampling. This chapter presents phylogenetic and chemotaxonomic characterisation of strain AFO1. The new isolate was found to be a genetically distinct strain of S. alaskensis. In light of the increasing number of isolated and characterised
Sphingomonas species and the recent sub division of the genus Sphingomonas into four distinct genera by Takeuchi et al. (2001), the taxonomic characteristics of S. alaskensis RB2256 are re-evaluated.
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2.2 MATERIALS AND METHODS
2.2.1 Bacterial strains and culture conditions
S. Alaskensis strains RB2256, RB2510 and RB2515 were obtained from Professor Jan Gottschal. Each strain was revived from stocks that were cryopreserved in 1992, transported on plates, immediately inoculated into artificial seawater medium (Eguchi et al., 1996), incubated overnight and preserved in 10% glycerol at -80˚C. Strain AFO1 was isolated from a depth of 350 m in the North Pacific, 1.5 km from Cape Muroto, Toyama Bay during early summer 1998 by Dr Mitsuru Eguchi. Strain AFO1 was brought onto plates only after 12 months of serial sub-culture in liquid filtered autoclaved seawater (FAS). S. alaskensis strains and strain AFO1 were maintained on ASW medium supplemented with glucose (3 mM), Sphingomonas macrogoltabidus ATCC51380 was maintained on R2A agar (Oxoid), Sphingomonas strain KT-1 (Tabata et al., 1999) was maintained on 3.0 g L-1 yeast extract, 5.0 g L-1 peptone and 0.75% NaCl (YEPS) and Halomonas subglaciescola ACAM 21 was maintained on Marine Broth 2216 (Difco). For preparation of DNA and fatty acids, S. alaskensis RB2256, strain AFO1 and H. subglaciescola were grown in Marine Broth 2216 (Difco), KT-1 in YEPS and S. marcrogoltabidus in R2Amediumat25°C, and cells harvested at late logarithmic phase (optical density 0.5 at 610 nm).
2.2.2 Alcohol precipitation
DNA was recovered from aqueous solutions by the addition of 0.1 volume of 3 M sodium acetate, pH 5.0, and 2.5 volumes of spectral grade ethanol. Samples were precipitated overnight at -15˚C or for 15 min at 0˚C. The DNA was then pelleted by centrifugation at 14000 x g for 30 min at 4˚C. After washing the pellet with cold 80% ethanol, the DNA was recentrifuged at 4˚C for 2 min, and air dried for 15 min.
2.2.3 Estimation of nucleic acid concentration
Nucleic acid concentrations were estimated spectrophotometrically using a Beckman Du 7500 spectrophotometer at wavelength 260 nm. Double stranded DNA 50 g 32 Chapter 2
ml-1 and single stranded DNA and RNA 40 g ml-1 corresponds to A260 of 1 (Sambrook et al., 1989). The purity of the DNA preparation was determined by the A260/A280 ratio. Solutions with an A260/A280 ratio of 1.8 or greater were considered to be pure. DNA concentrations were also estimated by comparing the relative intensities of DNA samples to standards of known concentration, electrophoresed on agarose gels.
2.2.4 Restriction enzyme digestion
Restriction enzyme digests were carried out in the buffer supplied for that enzyme by the manufacturer, or alternatively in One-Phor-All™ reaction buffer (Pharmacia). Digests contained 1-3 units of enzyme per g of DNA and were incubated for 1-2 h at the temperature recommended by the manufacturer.
2.2.5 Agarose gel electrophoresis and visualisation
DNA preparations such as restriction enzyme digests, PCR amplified DNA and plasmids were primarily analysed by electrophoresis on 0.7-1.5% (w/v) agarose (Progen DNA grade) gels. Agarose gel loading dye was added to samples (1:10 v/v) and electrophoresed in TAE buffer containing 0.5 g ml-1 EtBr at 80-120 mA for 1- 2 h (or overnight at 20 mA) on minigels (60 x 60 x 4 mm) or large gels (200 x 200 x 4 mm). Preparative agarose gels prepared aseptically with low melting point agarose (Sea Plaque ® GTG ® Agarose) (1.0 -1.5 % w/v in sterile E-buffer) were used to purify target DNA from contaminants. Gel forming apparatus was disinfected with 95% ethanol. Electrophoresis was performed at 4˚C in sterile TAE-buffer containing 0.5 g ml-1 EtBr. DNA was visualised using a long wave UV lamp. Section 2.5.2.4 outlines the isolation of DNA from preparative gels. DNA and RNA were visualised using a UV transilluminator at wavelength 254 nm and, when permanent records were required, photographs of gels were taken with a Mitsubishi video copy processor P67E.
33 Chapter 2
2.2.6 Genomic DNA extraction
Total genomic DNA was extracted using a modification of the method described by Murray and Thompson (1980). A 2 ml aliquot of late logarithmic phase cells was pelleted by centrifugation, the media decanted, and the pellet resuspended in 567 l of TE. Cells were lysed by the addition of 30 l of 10% w/v SDS and 3.0 l of 20 mg ml-1 proteinase K (Boehringer Mannheim) to give a final concentration of 100 g ml-1 proteinase K and 0.5% w/v SDS. The solution was mixed thoroughly and incubated at 60˚C for 4h before the addition of 100 l of 5 M NaCl and 80 l of 10% w/v CTAB (hexadecyltrimethyl ammonium bromide, Sigma) in 0.7% w/v NaCl. The CTAB/NaCl solution was prepared by slow addition of CTAB (10 g) to 100 ml of 0.7 M NaCl while heating and stirring.
Samples were mixed thoroughly and incubated at 65˚C for 10 min. CTAB complexes were extracted with 1 volume of chloroform:isoamyl alcohol (24:1 v/v) and centrifugation at 12 000 x g for 5 min and the supernatant was transferred to a fresh tube. Any CTAB complexes remaining in the supernatant were extracted with 1 volume of phenol:chloroform: isoamyl alcohol (25:24:1 v/v/v) and centrifugation at 12 000 x g for 5 min. The supernatant was transferred to a fresh tube and nucleic acids were precipitated by the addition of 0.6 volumes of isopropanol. After the tubes were mixed by gentle inversion the nucleic acids were collected by spooling on a glass rod and washed successively in 50%, 70% and 100% v/v ethanol. Spooled and washed DNA was transferred to a fresh tube, dried briefly in vacuuo and resuspended in deionised water. The DNA was subsequently used for 16S rRNA gene amplification, DNA-DNA hybridisation and mol% G+C content analysis.
2.2.7 Amplification of 16S rDNA sequences
Amplification of the almost full-length 16S rRNA gene from AFO1 was performed by PCR using the bacterial consensus 16S rRNA primers 27F and 1494R (Neilan et al., 1997). The amplification mixture (20 l) contained 25-50 ng of DNA, 0.2 mM
(each) deoxynucleoside triphosphates, 0.2 pM (each) primer and 2.5 mM MgCl2. After an initial denaturation step of 5 min at 95˚C, the temperature of the PCR mixture was lowered to 88˚C and 1-2 U Pfu Taq DNA polymerase (Sigma) was 34 Chapter 2
added. Thermal cycling was performed in a PCR Sprint Temperature Cycling System (Hybaid Ltd.) for 25 cycles of: 20 s denaturation at 95˚C, 20 s annealing at 50˚C, and extension at 72˚C for 1.5 min. The final extension step was 5 min at 72˚C followed by storage at 4˚C.
2.2.8 Purification and nucleotide sequencing of amplification products
PCR products were prepared for DNA sequencing by ethanol precipitation. After the addition of 0.1 volume of 3 M sodium acetate and 2 volumes of 80% v/v ethanol the tubes were vortexed for 20 s and incubated at 22˚C for 5 min. The DNA was collected by centrifugation at 12 000 x g for 10 min at 22˚C, the supernatant discarded, and the pellet resuspended in deionised water. DNA sequencing was performed using the PRISM BigDye cycle sequencing system (Applied Biosystems Inc.) and primers 27F, 519R, 530F, 929R, 1114F, 1221R and 1494R (Neilan et al., 1997) according to manufacturers instructions. Sequencing-reaction products were purified by ethanol precipitation, and sequence data was collected from a model 373 sequencer (Applied Biosystems). The DNA sequences were deposited in the GenBank database.
2.2.9 Phylogenetic analysis of 16S rDNA gene sequences
DNA sequences corresponding to Escherichia coli 16S rRNA gene positions 27 to 1433 were aligned using the programs Pileup, ClustalX (Thompson et al., 1997) and PHYLIP (Felsenstein, 1985). The nucleotide alignments were edited manually to resolve positions with ambiguities or gaps. The 16S rDNA distance trees were reconstructed using the neighbor-joining method with Jukes-Cantor corrections (Saitou and Nei., 1987) as implemented by ClustalX. The bootstrap confidence levels for the interior branches of the trees were estimated from 1,000 resamplings of the data (Felsenstein, 1985).
2.2.10 DNA-DNA hybridisation and mol%G+C content analysis
DNA-DNA hybridisation and mol%G+C content analysis were performed on DNA samples extracted at UNSW by Dr John Bowman at the University of Tasmania 35 Chapter 2
using methods adapted from Huss et al. (1983). Briefly, genomic DNA was sheared to an average size of 1 kb using sonication, dialysed overnight at 4°Cin2xSSC buffer (0.3 M NaCl, 0.03 M sodium citrate, pH 7.0), and adjusted in concentration to approximately 60-75 μgml-1. Following denaturation of the DNA samples, hybridisation was performed at the optimal temperature for renaturation (TOR)which was 25°C below the DNA melting temperature and was calculated from the following equation: TOR°C= 48.5 + (0.41 x %G+C). The decline in absorbance over a 40 min interval of DNA mixtures and control DNA samples were used to calculate DNA hybridisation values from the following equation: % DNA hybridisation = (4AB - A - B/2 (A x B)) x 100% A and B represent the change in absorbance for two DNA samples being compared and AB represents the change in absorbance for equimolar mixtures of A and B. DNA hybridisation values equal to or below 25% is considered to represent background hybridisation and are thus not considered significant.
The DNA base composition of strains was determined using the spectrophotometric thermal denaturation method described by Sly et al. (1986).
2.2.11 Fatty acid analysis.
Fatty acid analysis was performed on freeze-dried cell pellets by Dr. David Nichols at the University of Tasmania using the following methods. Lipids were extracted using the modified one-phase chloroform:methanol Bligh and Dyer extraction (Bligh and Dyer, 1959). A portion of the total lipid extract was transesterified by reaction at 80°C for 1 h using 3 ml of a methanol:chloroform:hydrochloric acid (10:1:1 v/v/v) solution. After the addition of water, the mixture was extracted with hexane:chloroform (4:1 v/v) to yield fatty acid methyl esters (FAME). Hydroxy functionalities were converted to OTMSi ethers by reaction with bis(trimethylsilyl)trifluoroacetamide (BSTFA) reagent at 80°C for 24 h.
FAME were analysed using a Hewlett Packard 5890 II gas chromatograph and 5970A Mass Selective Detector equipped with a 50 m x 0.22 mm internal diameter cross-linked methyl silicone (0.33 μm film thickness) fused-silica capillary column. 36 Chapter 2
Operating conditions are detailed in Nichols et al. (1986 and 1994). Identification FAME from all samples was achieved by interpretation of component spectra and comparison to those of known standards. Monounsaturated fatty acid double bond position and geometry was determined by GC-MS analysis of their dimethyl disulphide adducts (Nichols et al., 1994).
37 Chapter 2
2.3 RESULTS
2.3.1 Sequencing and phylogeny of strain AFO1
Almost full length (1,414 bp) sequence was obtained for the 16S ribosomal RNA gene (rDNA) of strain AFO1. The 16S rDNA sequence was compared with sequences in the GenBank sequence database and the most closely related sequences were identified by BLAST (http://www.ncbi.nlm.nih.gov/blast). The closest database matches were to two independently determined S. alaskensis RB2256 sequences,
Z73631 (Gottschal et al., submitted to GenBank 1996) displaying 1397/1413 identities, and AF148812 (Button et al., 1998) displaying 1369/1387 identities. High scoring matches were also obtained with S. alaskensis RB2256 co-isolates, strains
RB2515, RB2510 and RB255 (Vancanneyt et al., 2001). In light of the number of ambiguities and unassigned nucleotides in two database entries of S. alaskensis
RB2256 the 16S rDNA gene sequence of this strain was reanalysed using a stock culture cryopreserved in 1992. The 16S rDNA gene was amplified using Pfu high- fidelity DNA polymerase and unambiguous sequence of the rDNA gene was obtained for both DNA strands over a total of 1,414 bp, corresponsing to bases 27-
1441 of the Escherichia coli 16S rRNA sequence. The new sequence supercedes earlier incomplete entries, AF148812 and Z73631, which were obtained using less- stringent methods and error-prone, out-dated sequencing technology and should be used in place of them. The 16S rDNA sequences for strains RB2256 and AFO1 were deposited in the GenBank database under accession number AF378795 and
AF378796 respectively.
38 Chapter 2
The determined sequences for AFO1 and RB2256 along with related sequences from
GenBank were used for constructing phylogenetic trees. Tree topology was similar for parsimony and distance matrix trees and a representative neighbour-joining tree is shown (Fig. 2.1). The marine S. alaskensis strains along with strain AFO1 form a mono-phyletic cluster indicating that strain AFO1 is a strain of S. alaskensis.From the alignments used for tree construction, only one nucleotide change was present between strain AFO1 and RB2256 (1,414 bp) (Table 2.1). S. alaskensis strains
RB2510 and 2515, that were isolated from Resurrection Bay (Vancanneyt et al.,
2001), had one and two nucleotide changes respectively and four alignment gaps compared with the sequence for strain RB2256. In view of the errors for 16S rRNA sequences for strain RB2256 (accession numbers Z73631 and AF148812), these differences may also be sequencing artefacts. The most similar sequence to those in the S. alaskensis cluster is from strain KT-1 (Fig. 2.1, Table 2.1) which has 19 nucleotide differences. The sequence of a Sphingomonas isolated from a hot-spring
(acc. AB015049) was also very similar to AFO1 (98.1% identity, 1311 identities
/1338 residues, no gaps).
39 Chapter 2
TABLE 2.1. Changes and gaps relative to S. alaskensis RB2256T 16S rDNA sequence
Strain Differences1 gaps/insertions1 AF01 (AF378795) 1 0 RB2510 (AF145753) 0 4 RB2515 (AF145754) 1 4 KT-1 (AB022601) 18 1 MBIC3365 (AB015049) 272 02 1 nucleotide changes relative to 1414 bp of RB2256T 16S rDNA sequence (AF378796) 2 nucleotide changes relative to 1338 bp alignment with RB2256T 16S rDNA sequence.
40 Chapter 2
FIGURE. 2.1 Distance-matrix tree of selected Sphingomonas 16S rDNA sequences. DNA sequences corresponding to the E. coli 16S rRNA gene positions 27 to 1433 were aligned using the programs PILEUP and ClustalX (Thompson et al., 1997). Genetic distances were calculated using the method of Jukes and Cantor, and the phylogenetic tree was reconstructed using the neighbor-joining algorithm of Saitou and Nei (1987) as implemented within ClustalX. The phylogenetic tree was plotted using the program nj-plot. The root of the tree was determined using the 16S rDNA gene of Bacillus (BCE277907) as an outgroup. Numbers on branches represent bootstrap values for 1000 repeats (Felsenstein, 1985). The bar indicates 2 nucleotide changes per 100. Accession numbers unless indicated on the figure: (1) AF145754, (2) AF378795, (3) AF378796, (4) AF145753, (5) AB022601, (6) AB015049, (7) AF367204, (8) AF181572, (9) D13723, (10) AF327069, (11) AY081981, (12) D17322, (13) D13727, (14) U20756, (15) X94102, (16) D16144, (17) AF125194, (18) M96746, (19) AF510191 and (20) AF327028.
41 Chapter 2
2.3.2 DNA base composition and DNA-DNA hybridisation
The high degree of sequence identity between strain AFO1 and RB2256 raises the possibility that these two strains are genetically identical. In order to assess the extent of identity between these strains at a high level of resolution the mol% G+C content was determined and DNA-DNA hybridisation was performed. DNA from
Sphingomonas sp. strain KT-1 was used as an external reference. The mol%G+C for
DNA from strains RB2256, AFO1 and KT-1 were 64.8, 65.1 and 65.3, respectively
(Table 2.2). The DNA-DNA reassociation level between genomic DNA from strains
RB2256 and AFO1 averaged 84% while between strains RB2256 and KT-1 DNA hybridisation levels averaged only 34% (Table 2.2). S. macrogoltabidus ATCC
51380 (mol% G+C 65) and H. subglaciescola ACAM 21 were used as control strains
(mol%G+C 59) and both exhibited background renaturation levels of 12 to 23% with strain RB2256. S. macrogoltabidus exhibited significant levels of hybridisation
(44%) with strain KT-1 however below the level indicative of a bacterial species
(Stackebrandt and Goebel, 1994,Wayne et al., 1987). This indicates that strain KT-1 is closely allied to the species S. macrogoltabidus but appears to represent a novel species.
TABLE 2.2 DNA base composition and DNA:DNA hybridisation values between S. alaskensis RB2256, strains AFO1and KT-1, and S. macrogoltabidus.
Strain Mol% G+C (Tm) RB2556 KT-1 %DNA hybridisation S. alaskensis RB2256T 64.8 100 AFO1 65.1 84 KT-1 65.3 34 100 S. macrogoltabidus 65.4 23 44 ATCC51380
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2.3.3 Fatty acids
The fatty acid composition of Sphingomonas sp. KT-1 and AFO1 compared to S. alaskensis RB2256 is shown in Table 2.3. S. alaskensis RB2256 and strain AFO1 were grown in ASW supplemented with 3 mM glucose. Sphingomonas strain KT-1 was grown in tap water supplemented with 3.0 g l-1 yeast extract, 5.0 g l-1 peptone,
0.75% w/v NaCl. Samples were taken at mid-exponential growth phase and the washed cell pellets were freeze-dried before fatty acid and methyl ester analysis
(FAME). FAME analysis was carried out by Dr. David Nichols. The two marine strains (AFO1 and S. alaskensis RB2256) possessed a similar fatty acid profile,
(major components being 17:1w6c, 18:1w7c and 17:1w8c) and were distinguished from the freshwater species KT-1 (containing 18:1w7c and 16:1w7c as major components). The fatty acid profiles determined support the phylogenetic relationship between KT-1, AFO1 and S. alaskensis RB2256 as described by 16S rRNA sequence (Fig. 2.1).
43 Chapter 2
TABLE 2.3 FAME analysis of fatty acids from S. alaskensis RB2256 and Sphingomonas strains AFO1 and KT-1.
Percentage composition1 Fatty Acid KT-1 AFO1 RB2256
14:0 0.5 ± tr - - - - 15:0 0.2 ± tr 1.9 ± 0.1 2.0 ±tr 16:0 3.4 ± tr 3.6 ± 0.2 3.2 ±tr 17:0 - - 3.2 ± 0.2 4.0 ±tr 18:0 tr ± tr 0.4 ± tr 0.9 ±1.0 Sum Saturates: 4.1 ± tr 9.1 ± 0.4 10.1 ±1.0
i15:0 - - - - 0.3 ±tr Sum Branched: - - - - 0.3 ±tr
14:1w5c 0.4 ± tr - - - - 15:1w6c - - 0.2 ± tr 0.2 ±tr 16:1w7c 34.2 ± 0.2 4.3 ± 0.2 3.8 ±tr 16:1w5c 3.1 ± tr 0.8 ± tr 0.7 ±tr 17:1w8c 0.3 ± tr 8.1 ± 0.5 10.0 ±0.5 17:1w6c 1.9 ± tr 35.8 ± 1.8 44.7 ±0.3 18:1w7c 42.6 ± tr 24.3 ± 1.2 20.1 ±0.1 18:1w5c 1.2 ± tr 3.8 ± 4.8 0.9 ±tr 19:1w8c - - 0.9 ± tr 1.0 ±0.1 19:1w6c - - 1.2 ± 0.1 1.3 ±0.2 Sum Monounsaturates: 83.6 ± 0.2 79.3 ± 1.0 82.7 ±0.1
2-OH14:0 3.8 ± tr 1.2 ± 0.1 0.6 ±tr 2-OH15:0 - - 5.1 ± 0.3 3.5 ±tr 2-OH16:1w5c 0.3 ± tr 0.4 ± tr 0.1 ±0.1 2-OH16:0 7.6 ± 0.1 3.1 ± 0.2 1.5 ±1.0 2-OH17:1w6c 0.3 ± tr 1.8 ± 0.1 1.2 ±0.2 2-OH18:1w7c 0.4 ± tr - - tr ±tr Sum Hydroxy: 12.4 ± 0.2 11.6 ± 0.6 6.9 ±1.1
Total 100.0 ± tr 100.0 ± tr 100.0 ±tr 1: number of replicates (n) = 3; data presented as average ± standard deviation; tr = trace proportion, defined as less than 0.1%.
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2.4 DISCUSSION
2.4.1 Ecological implications of S. alaskensis sp. strain AFO1
The 16S rDNA sequence, mol%G+C, DNA-DNA hybridisation and fatty acid composition indicate that strain AFO1 is almost identical to S. alaskensis RB2256.
The level of sequence identity between strain AFO1 and RB2256 is striking. A result that suggests that these two strains, isolated from the North Pacific near Japan and Alaskan waters respectively, over a period spanning 10 years, are derived from a common ancestor in recent history. Pacific Ocean currents may account for the dispersion of S. alaskensis the 10, 000 km from Alaskan waters to Japan, or vice versa, for example, the North Pacific Intermediate Water current moves water through locations near these two sites (You et al., 2000). The presence of this single species as a numerically significant proportion of the bacteria at geographically remote sampling sites demonstrates the ability of S. alaskensis to proliferate in diverse ocean environments, from the permanently cold waters (4˚C) of Ressurection
Bay (Schut et al., 1993) to the relatively temperate water of the Pacific near Japan.
Strains AFO1 and RB2256 were also isolated from different depths (350 m and 10 m respectively) demonstrating that this species is capable of competing in distinctly different niches within the water column.
In addition to the type strain, S. alaskensis RB2256, and at least six other strains that were isolated from Resurrection Bay in Alaska (Schut et al., 1997a, Vancanneyt et al., 2001), morphologically similar strains have been isolated from the North Sea
(Schut et al., 1997a). A range of distantly related Sphingomonas strains have been isolated from oligotrophic and eutrophic marine environments including the Baltic
45 Chapter 2
Sea (Pinhassi et al., 1997, Pinhassi and Hagstrom, 2000), the North Sea (Eilers et al.,
2000), below sea ice off eastern Antarctica (Bowman et al., 1997) and as pathogens of corals (Richardson et al., 1998). Estimates of the abundance of Sphingomonas strains in marine environments have been reported by Pinhassi et al. (1997) in the
Baltic sea, (e.g. Bal 46 and BAL35, 18.9% and 2.4% of total bacterial numbers respectively) and by Schut et al. (1993) who isolated Sphingomonas strains with abundances of 15 to 35%.
The fact that two isolates, separated by more than 10,000 km can have almost identical sequences is intriguing, especially when the RB2256 co-isolates, which display less sequence identity at the level of 16S rDNA, are considered. DNA-DNA hybridisation provides some resolution to the relationship between strains AFO1 and
RB2256. The level of genomic DNA-DNA hybridisation (84%) indicates that strain
AFO1 displays significant differences at the genomic level and is therefore a genetically distinct strain of S. alaskensis. Such differences could be the result of genomic rearrangement, deletions, integration of phage or other mobile genetic elements. In contrast, strains RB255, RB2510 and RB2515 display greater similarity to RB2256 (> 89%) as determined by DNA-DNA hybridisation (Vancanneyt et al.,
2001).
The fatty acid profiles determined support the phylogentic relationship between KT-
1, AFO1 and S. alaskensis as described by the 16S rDNA sequence. Species of all four Sphingomonodaceae genera described by Takeuchi et al. (2001) are characterised by the presence of 2-OH14:0. In addition to this component a series of further 2-OH fatty acids were identified from strains KT-1, AFO1 and RB2256 by
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FAME analysis. There is a noticeably higher ratio of 17:1/18:1 and 16:1/16:0 fatty acids in S. alaskensis strains as compared with those from the Sphingomonas,
Novosphingobium and Sphingobium clusters. This general feature is found in other members of the Sphingopyxis genus (Takeuchi et al., 2001) and may serve to distinguish members of this genus from the other genera. It is interesting to note the general quantitative differences for fatty acid groups presented here in comparison with data previously reported for S. alaskensis (Vancanneyt et al., 2001). Together, with the low ratio of 17:1/18:1 and 16:1/16:0 fatty acids measured for the phylogenetically closely related freshwater species, strain KT-1 (1394/1414 identities in 16S rDNA sequence, 98.6%, Table 2.3), these differences highlight the dependence of fatty acid profiles on growth conditions, e.g. high NaCl vs no NaCl, media richness and growth phase.
2.4.2 Reclassification of Sphingomonas alaskensis to Sphingopyxis alaskensis
Phylogenetic analysis of existing Sphingomonas 16S rDNA gene sequences indicate that the currently known members of the genus could be resolved into four distinct clusters (Takeuchi et al., 2001). In support of these, distinctions based on gene sequencing, chemotaxonomic and phenotypic differences were noted between each cluster. Three new genera were proposed in addition to the genus Sphingomonas sensu stricto. According to this reclassification scheme S. alaskensis strains RB2256,
AFO1 and related strains belong to the genus Sphingopyxis with Sphingopyxis macrogoltabidus IFO 15033T (formerly Sphingomonas macrogoltabidus) as type strain. Characteristics that were used for distinguishing Sphingopyxis from the other
‘Sphingomonas’ clusters include greater than 97% 16S rDNA identity to the type
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strain, the absence of nitrate reduction, the accumulation of the polyamine spermidine rather than homospermidine (Takeuchi et al., 2001), and the fatty acid profiles as outlined in the results and the discussion below. It is notable that individual members of each cluster have been isolated from diverse environments and that the 16S rDNA delineation does not support groupings based on the habitat from which they were isolated. This is reflected by the diverse range of environmental conditions from which members of the Sphingomonodaceae have been isolated (Takeuchi et al., 2001 and references therein). Unusual environments from which Sphingomonads have been isolated include the accretion ice above Lake
Vostok (Christner et al., 2001), the deep subsurface (Balkwill et al., 1997), activated sludge (Neef et al., 1999) and hydrocarbon polluted soils (Baraniecki et al., 2002). It is interesting to note the high degree of 16S rDNA sequence similarity of strains
RB2256 and AFO1 to strains KT-1 and MBIC3365 (Table 2.1) considering that KT-
1 was isolated from natural mineral water and strain MBIC3365 was isolated from a halophilic spa. As evidenced by their wide distributions and metabolic diversity members of the family Sphingomonodaceae appear to be generalists, capable of adapting to a wide range of physical and biochemical environments.
2.4.3 Conclusion
Strain AFO1 is almost identical to S. alaskensis RB2256. Work done by colleagues has established that morphology, genome size and stress resistance profiles of strain
AFO1 are identical to S. alaskensis RB2256. This collective work was published by
Eguchi et al. in 2001. The isolation of AFO1 indicates a broad geographical distribution, persistence and environmental significance for S. alaskensis genotypes.
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In light of the overall level of similarity between AFO1 and the type strain RB2256, only RB2256 was used for the remainder of studies performed in the following chapters.
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3CONTINUOUS CULTURE METHODS FOR
INVESTIGATING PHYSIOLOGICAL
RESPONSES TO NUTRIENT LIMITATION IN
MARINE BACTERIA
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3.1 BACKGROUND
In order to comprehend microbiological processes and the environmental processes in which they participate, it is essential to understand the physiology of microorganisms with respect to the prevailing environmental conditions. In the oligotrophic ocean the limited availability of nutrients limits microbial growth and has a major impact on microbial physiology. Growth limitation relates largely to the availability of utilisable nutrients, such as carbon, nitrogen and phosphorus, however, the availability of trace metals, vitamins and a variety of physicochemical factors may also impact on the abundance of microorganisms.
Organisms in oligotrophic environments inhabit a constant limitation. Although simple laboratory systems cannot capture the richness and complexity of natural ecosystems, such systems do permit careful measurement of cell physiology under well-defined and reproducible conditions. Furthermore, well-defined laboratory systems offer a starting point for studying responses to controlled environmental changes and allow the rigorous experimental testing of hypotheses (e.g. Gottschal,
1990a, Velicer et al., 1999). The traditional and perhaps most straight-forward method of microbial cultivation is the batch culture. Within this system initially all nutrients are in excess allowing the organism to grow at an optimal, unlimited rate.
At a point where the concentration of one or more nutrients become limiting the exponential growth rate diminishes and eventually ceases and the organism enters a starvation state. The main disadvantage of this system is that exponential growth and near zero growth in stationary phase are the growth modes that may be studied in detail. The intervening phases of decelerating growth, corresponding to the period of
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nutrient limitation, rapidly merge into one another and are therefore difficult to study in isolation.
While still poorly studied, there is expanding interest in the effects of growth rate, as may be imposed by nutrient limitation in chemostats, on the physiology of microorganisms. For example, cell size, cellular composition and starvation survival are affected by growth rate in Vibrio sp. strain ANT-300 (Moyer and Morita, 1989a,
1989b), and slow growth regulates porin expression and induces cAMP- and RpoS- dependent gene expression in E. coli (Matin and Matin, 1982, Schultz et al., 1988,
Notley and Ferenci, 1995, 1996, Liu and Ferenci, 1998, Tweeddale et al., 1998).
Other regulatory mechanisms that respond to the level of nutrient limitation in E. coli, such as the stringent response and regulation of mutation rates themselves, have been identified (Cashel et al., 1996, Notley-McRobb et al., 1997, Ferenci, 1999).
In response to starvation copiotrophic bacteria become resistant to a variety of stresses (e.g. Matin, 1990, Nyström et al., 1990ab, Nelson et al., 1997).
The chemostat is a basic piece of laboratory apparatus that allows the study of bacteria growing at sub-optimal rates. Chemostat studies have begun to occupy an increasingly central role in ecological and evolutionary studies (e.g. Velicer et al.,
1999, Notley-McRobb and Ferenci, 1999, Stahl et al., 2004). The environment created by a chemostat is one of the few completely controlled experimental systems for testing microbial growth and competition. The theory and procedures for continuous cultivation have been reviewed extensively (Gottschal, 1990a and
1990b). A basic diagram of a chemostat set-up, comparing the two types of chemostats employed in this study is presented in Figure 3.1. The basic parameters
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defining chemostat growth are presented in Table 3.1. In the chemostat, the cell density and culture volume are kept constant and the growth rate of the culture is fixed externally by the rate at which fresh medium is supplied. After a period of equilibration the specific growth rate ( ) is considered to be equal to the dilution rate
(D):